Special thanks to Dr. Clare M. Waterman for mentorship and discussions, Michelle Baird and Mike Davidson for providing many of the fluorescent cDNA constructs used in these studies, and Kathryn Applegate and Gaudenz Danuser for developing plusTipTracker MT analysis software. This work was supported, in part, by a grant to K.A.M from the National Heart Lung and Blood Institute (NHLBI) and the National Institutes of Health (NIH). (Grant: 4K22HL113069).

The authors declare that they have no competing financial interests.

Using HUVECs in Morphology and Migration Experiments. 
Live-cell imaging of EB3-labeled MT growth dynamics within HUVECs requires appropriate and reproducible cell culture conditions in order to measure and evaluate changes in MT growth and HUVEC morphology, such that they can then be assigned to an ECM signaling cue. HUVECs represent an excellent model for studying cell shape and motility. They are highly amenable to transfection with fluorescently-labeled cDNAs, they exhibit dramatic morphologies in response to both soluble and physical signaling cues, and they maintain the ability to organize themselves into vascular tubes when provided appropriate cell culture conditions65-70. Primary cell cultures, such as HUVECs, require careful handling and monitoring of cell passage number in order to ensure the cells are healthy and that they will behave normally during experimentation. For example, when conducting experiments investigating the cytoskeleton and/or cell morphology, HUVECs should not be used for experiments beyond the tenth cell culture passage, as HUVECs typically begin to display non-uniform morphologies around passage twelve to fifteen.
Cell density is also a critical regulator of HUVEC health and proliferation. HUVEC branching investigations (see Protocol 10.1-10.4, above) use relatively low cell density cultures in order to provide the cells physical space to interact with their ECM and to extend branched protrusions. In contrast, in wound-edge migration studies (see Protocol 10.5-10.8), HUVECs are culture in a monolayer prior to wounding. As such, wound-edge HUVECs display a relatively un-branched morphology, extend leading edge protrusions, and exhibit cell-cell interactions with their immediate neighbors as they migrate into the wound.
Caveats and Advantages of Tracking MT Dynamics in Living HUVECs. 
Spinning disk microscopy is an excellent approach to test experimental hypotheses that require confocal image clarity with high time-resolution. With the appropriate camera and laser module, this microscopy approach is sensitive to low emission fluorescence and can aid in reduced photobleaching compared to other types of fluorescence microscopy. Importantly, this approach is also well suited for moderate to high time-resolution imaging as is required for generating comprehensive measurements of MT growth dynamics using the EB3 labeling approach, in a variety of different cell types.
While the EB3 methodology of labeling growing MT plus-ends has distinct advantages over other methods of fluorescently labeling MTs, it also has unique disadvantages. For example, there is a proportion of the MT array that is not actively growing at any given time, including both the population of disassembling MTs and the population of stabile, non-dynamic MTs71-74. EB3 would not label these two populations of MTs and therefore the contribution of these two populations would not be included in experimental data analysis. Alternative methodologies for evaluating the entire MT array have focused on expression of fluorescently-labeled tubulin, which incorporates into all populations of MTs and allows for identification of growing, shrinking, and stabile MTs. However, because of the relatively high density of the MT array near the center of the cell and adjacent to the microtubule organizing center (MTOC), labeling the entire MT array creates a disadvantage in performing image analysis of MT ends that are not near the cell periphery75-77. Experiments using two-color co-labeling of MTs with fluorescent tubulin and fluorescent EB3 in live cells have revealed, qualitatively, that the vast majority of tubulin-labeled MTs are also EB3-labeled78. Additionally, there has not been an efficient method of automated tracking of MTs labeled with fluorescent tubulin. This means that data collection and measurements of MT dynamics in cells expressing fluorescent tubulin require hand tracking of individual MTs, a process that is error prone and tedious. As a result, automated computational analysis of EB3 labeled MTs has become the preferred methodology for measuring MT dynamics as it can be applied to MT dynamics experiments across numerous cell types and within a variety of experimental regimes35,79.
Linking MT Growth Dynamics to HUVEC Morphology and Migration. 
From the perspective of a cytoskeletal investigator, one extremely useful adaptation of the plusTipTracker automated tracking analysis is the ability to measure and evaluate both whole cell and regional changes in MT growth dynamics. The requirement for a cell to become polarized is an essential pre-requisite for directional cell migration. As such, polarized signaling cues have long been known as key driving factors for directional cell migration30,31,51,54,56,80,81. For example, soluble signaling molecules such as VEGF induce actin polymerization and MT growth toward the VEGF cue in endothelial cell types2,82-87. Additionally, in response to physical interactions with the ECM (for example, compliance and dimensionality mechanosensing), HUVECs modulate their MT dynamics by locally regulating MAPs to promote MT growth within elongating branched protrusions, and this serves to guide HUVEC migration in the direction of the cue5,31,88. Thus, polarization in response to extracellular signaling cues highlights the need for experimental evaluation of localized changes in cytoskeletal dynamics.
Simultaneous live-cell imaging of MT growth dynamics (using the EB3 approach) and HUVEC cell migration is extremely difficult to perform because MTs grow on a time scale of seconds, while changes in cell morphology and directional migration occur on a time scale of hours. Yet both processes are intimately linked. Moreover, continuous, rapid imaging of fluorescently-labeled EB3 on the second time scale leads to photobleaching of the EB3 signal. Attempts to overcome this issue by conducting short imaging series of EB3 (1-2 min acquisition series) every 30 minutes have been successful, but could only be accomplished in a small number of ECs that had optimal expression of EB3 and that did not display adverse effects to repeated laser illumination5.
An alternative approach that allows for interpretations of how MT growth dynamics contribute to changes in cell morphology and migration is the plusTipTracker Region of Interest (ROI) tool35 (see Protocol 6.6). This methodology allows for whole cell MT growth tracks to be sub-divided into user-defined regions, from which MT growth tracks can then be extracted and analyzed. For example, comparisons of &#34;branched&#34; versus &#34;non-branched&#34; regions of the cell have revealed that MTs grow more slowly and more persistently within branched regions as compared to the non-branched cell periphery5. In polarized wound-edge HUVECs, comparisons of the leading edge and trailing edge MT growth dynamics revealed that MCAK-induced MT disassembly is inhibited specifically within the leading edge, but not the trailing edge. Regional evaluation of leading and trailing edge MT growth dynamics in HUVECs expressing Rac1 and/or phosphorylation mutants of MCAK further revealed that Rac1-induced activation of Aurora-A kinase specifically and locally inhibits MCAK activity within the leading edge to drive HUVEC polarization and directional migration31. Thus, the combination of automated tracking of MT growth dynamics and regional evaluation of MT growth dynamics within the branched protrusions and/or the leading edge of wound-edge HUVECs has allowed us to link MT growth dynamics to HUVEC morphology and migration.
The protocols described are designed to provide a generally straightforward and highly repeatable methodology for HUVEC culture and experimental investigation. Nevertheless, many of the protocol details are complex and time consuming, which, in turn, provides opportunity for errors or difficulties in conducting the experimental methodology. While it was attempted to address potential problems throughout the protocol, here is provided additional, detailed troubleshooting of potential problems that are less common or otherwise abbreviated as notes within the methodology protocols.
Related to coating coverslips with polyacrylamide substrates (Protocol 2.2), a common problem that arises is the complex nature of activating glass coverslips, coupling polyacrylamide to the glass, activating polyacrylamide, and then coupling ECM to the polyacrylamide. One particularly difficult and potentially problematic step is culturing HUVECs onto the fibronectin/polyacrylamide gel following exposure of the polyacrylamide-coated coverslips to 2 mM sulfo-SANPAH (step 2.2.2.6, above). It is critical to the success of culturing HUVECs on these substrates that sulfo-SANPAH be completely removed from the experimental system following ECM conjugation. This can be best accomplished by washing with ddH2O and incubating the coverslips in ddH2O on a shaker overnight. If cells cultured on the polyacrylamide ECMs do not survive, or if viability is less than expected, increased and repeated washing of sulfo-SANPAH after conjugation to fibronectin (step 2.2.2.9) is recommended to potentially alleviate this problem.
Related to Protocol 3 (cDNA transfection and HUVEC culture on a coverslip), a major influence to the success of the electroporation procedure is the handling of the HUVECs both during trypsinization of the cells to remove them from the plastic flask, and following the conclusion of the electroporation (steps 3.3-3.4, above). It is critical to carefully monitor the HUVECs while in trypsin, and not to exceed the time needed for the cells to &#34;round-up&#34; on the surface of the flask (typically 1-2 min). Excessive time in trypsin is a major cause of HUVEC death, which is typically not realized until the cells are plated onto fibronectin-coated coverslips (step 3.4) and then fail to attach to the substrate.
Of similar importance, HUVECs (and most primary cell types) do not fare well when suspended for long times in the electroporation buffer. During larger-scale experiments, for example, where the user has five or more conditions that require electroporation, it is better to keep the cells pelleted in individual tubes (1 tube per transfection) and to mix them, one-at-a-time, in the electroporation buffer immediately before electroporation. This approach minimizes incubation time in electroporation buffer and maximized HUVEC survival. Additionally, immediate rescue in complete culture media is also critical following electroporation. Rescue delays of one minute or more following electroporation can result in near complete HUVEC death and thus should be avoided.
Related cell migration assay (Protocol 9), the technique of culturing HUVECs at extremely high density depends on a critical modification (described in step 9.2) of the traditional HUVEC cell culture methodology (described in Protocol 3) that requires drying of the fibronectin coated coverslip in order to enable HUVEC culture into a very small area of the coverslip. Often times, if the coverslip is not completely dried or if the prior coating of the coverslip with fibronectin (steps 2.1 or 2.2) was not efficient, the media-containing cells that is placed on the coverslip will lose its surface tension and expand to the edges of the coverslip, thereby reducing the density of cells on the coverslip. One method to troubleshoot this potential problem is to aspirate the media from the coverslip and then place the coverslip (still in its 35mm dish) into the back of the biosafety cabinet for 5-10 min. This adaptation can help to allow the flow of air within the cabinet to assist in drying the dish. If any visible liquid remains on the coverslip after air drying, aspirate the liquid spots and again allow to air dry for 5-10 additional min in the biosafety cabinet. It is important to note that there is no way to completely avoid this potential problem, but it does become less of an issue with repeated practice of the protocol. It is also a troubleshooting recommendation that when preparing coverslips for the cell migration assay (or any assay, in general), to prepare extra coverslips and have extra HUVECs ready-to-go in case of problems along the way.
With these troubleshooting considerations in mind, it is also important to highlight the utility of the protocols discussed and their implications in future cell biology research. The applicability to the polyacrylamide approach is wide-ranging, and includes not only two-dimensional studies of cell-ECM engagement (described above), but also three-dimensional investigations5. This application is critical to increasing our understanding of how HUVECs regulate cell shape and migration in physiological environments and should provide investigators a basic understanding of how morphological changes are coordinated with cytoskeletal changes. While the protocols described here are focused on measurements of MT growth dynamics, the majority of the protocols can and should be adapted to measure any number of other microscopically measureable aspects of cell-ECM interactions. With respect to MT dynamics, there are numerous MAPs, including at least 14 families of kinesins89, each with a potential role in contributing to HUVEC polarity and directed migration, and all of which can be investigated using the protocols presented.
Future investigations should also be targeted at cell-ECM engagement in normal versus diseased states. HUVEC protocols presented here are applicable to investigations of normal vascular homeostatic processes, as well as to disease states including heart disease, retinopathy, tumor growth and metastasis, to name a few. Conjugating visibly transparent ECMs and polyacrylamide substrates of amenable compliance with microscopic studies will allow for cell-ECM engagement studies such that endothelial cell vascular angiogenesis can be monitored with high spatial and temporal resolution. These types of experimental approaches have already begun to reveal important differences in endothelial cell shape, polarity, migration, and the effects of proteins that regulate these processes5,31,90. Future efforts should aim to identify how the combination of ECM compliance and dimensionality mechanosensing may serve to locally control proteins that target and regulate cytoskeletal dynamics.

Angiogenesis is triggered by extracellular signaling cues that promote endothelial cells (ECs) to polarize, migrate with directional specificity, and form new vasculature in tissues that require blood supply. Contributing factors thought to drive angiogenesis are signals from the extracellular matrix (ECM). In response to physical cues, ECs extend new branches with directional specificity to guide directional movement in formation of the vascular network1,2. The architecture of EC morphology and branching is defined by localized regulation of the microtubule (MT) cytoskeleton in a way that is coordinated with the regulation of acto-myosin contractility3. In order for EC branches to form, myosin-II must be locally downregulated, resulting in localized membrane protrusions4. At the same time and in the same location within the cell, MT growth dynamics become modified resulting in slow and persistent MT growth to promote branch elongation and stability5. This coordinated cytoskeletal regulation allows ECs to polarize their morphology to facilitate directional migration.
The development of a polarized cell morphology requires polarized MT assembly6. In migrating epithelial cells, MTs grow slowly and persistently specifically at the leading edge of the cell7-9; while in neurons, the initiation and elongation of axons or dendrites requires that MTs are not only present, but that they are dynamically undergoing the processes of assembly and disassembly10-15. In this way, localized control of MT growth dynamics determines the architecture of the cell and allows for rapid remodeling of cell morphology as ECs navigate through their ECM.
MT growth dynamics are regulated by families of molecular motor proteins and non-motor proteins, termed Microtubule Associated Proteins or Microtubule Interacting Proteins (henceforth referred to as MAPs). MAPs can be locally activated or inactivated, typically through signaling cascades initiated at the cell-ECM interface16,17. Once active, MAPs function by binding to the MT and altering the kinetics of MT assembly and disassembly; thereby functioning to locally regulate MT dynamics in order to promote cell shape and polarity18-20. Mitotic Centromere Associated Kinesin (MCAK) is a Kinesin-13 family MAP that binds to MT ends and couples ATP hydrolysis to MT disassembly21-23. This functional role of MCAK is known to be critical within the mitotic spindle24-28, as well as during interphase29,30. Within interphase ECs, MCAK-mediated MT disassembly is regulated by localized Aurora-A induced phosphorylation of MCAK in response to wound-edge induced activation of the Rac1 small GTPase. The outcome of this signaling cascade is the inhibition of MCAK at the leading edge of ECs, resulting in polarized MT growth toward the leading edge and MCAK-induced MT disassembly at the trailing edge of migrating ECs31.
Traditional methodologies for measuring MT growth dynamics have typically used hand tracking of either fluorescently-labeled tubulin or, more recently, fluorescently-labeled MT End Binding protein 1 or 3 (EB1 or EB3, respectively). The fluorescent tubulin approach has the advantage of labeling the entire MT array. However, because the majority of mitotic cell types maintain a radial array of MTs during interphase8,32,33, MTs near the centrosome are very dense, and as a result, tracking MT growth and disassembly with fluorescent tubulin is usually limited to the peripheral regions of the cell. Because of these limitations, utilization of fluorescently-labeled EB proteins (EB1 or EB3) has been the more common approach to measuring MT dynamics. Because EBs only label the growing plus-ends of MTs, they appear as small (1-3 &#956;m), brightly fluorescent comets, thus providing an easily discernable marker of MT polymerization with very limited background fluorescence34-37. While the fact that EBs label only a subset of the MT array represents a major strength of the EB approach, it also highlights one important limitation, that non-growing MTs are not labeled by the EB3 approach. Nevertheless, the ability to measure and track EB3 comets over time and to visualize MT growth within sub-cellular regions make this approach ideal for applications that require regional evaluation of MT organization.
Another critical limitation of traditional methodologies for measuring MT growth dynamics has been very large data sets and the time required for data analysis. This limitation stems from the need for hand tracking of hundreds of EB comets at high time resolution. Moreover, these measurements need to be done in a statistically significant number of cells and also performed under a variety of control and experimental conditions. Recently, these constraints have been overcome through computational analysis techniques specifically directed at tracking EB3 comets using time-lapse microscopy in living cells35. When used appropriately, this computational approach serves to both restrict the potential for human error associated with hand-tracking in addition to providing for a high-throughput method of measuring MT polymerization. Computational analysis of MT dynamics using the MatLab-based software package, plusTipTracker, allows for measurements of MT growth by tracking fluorescently-labeled EB3 comets5,31,35,38. Additionally, plusTipTracker software allows for calculations of MT catastrophe events (a.k.a. disassembly) based on the disappearance of EB3 comets within a growing MT track, and allows for sub-cellular (a.k.a. regional) analysis of MT growth dynamics within user-defined regions of interest. For our studies, MT growth dynamics were compared within branched regions versus non-branched regions, or within leading versus trailing cell edges. Data output from plusTipTracker software can also be used to generate color-coded track overlays for individual cells and sub-cellular regions.
Here we describe the methodology used to investigate the role of MCAK in modulating MT growth dynamics and the contribution of this MT depolymerase to the regulation of EC branching morphology and directional migration. Our approach used a primary human cell line, Human Umbilical Vein Endothelial Cells (HUVECs), which are easily cultured and highly amenable to electroporation-induced, transient transfection of plasmid cDNA. We then used high-resolution, time-lapse fluorescence microscopy of HUVECs transfected with MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics transfected HUVECs. PlusTipTracker-based computational image analysis of EB3-labeled MT plus ends was to measure MT growth speeds and MT growth lifetimes in time-lapse images, to measure and compare effects of experimental manipulations on MT growth dynamics. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration. These techniques are applicable to investigations of any MAP that can be fluorescently labeled and expressed in live cells.

<table><tbody><tr><td>Aurora A Inhibitor I</td><td>Selleck Chemicals</td><td>S1451</td><td></td></tr><tr><td>2% Bis solution</td><td>Bio-Rad</td><td>1610142</td><td></td></tr><tr><td>3-aminopropyltrimethyloxysilane</td><td>Sigma-Aldrich</td><td>A3648</td><td></td></tr><tr><td>40% acrylamide</td><td>Bio-Rad</td><td>1610140</td><td></td></tr><tr><td>405, 488, 561, 640 nm high power MLC400 laser combiner module</td><td>Agilent Technologies</td><td>99462</td><td></td></tr><tr><td>Ammonium persulfate</td><td>Sigma-Aldrich</td><td>A3678</td><td></td></tr><tr><td>anti&amp;ndash;mouse IgG immunoprecipitation beads</td><td>TrueBlot Reagents; Rockland Inc.</td><td>88-1688-31</td><td></td></tr><tr><td>anti&amp;ndash;rabbit IgG immunoprecipitation beads</td><td>TrueBlot Reagents; Rockland Inc.</td><td>88-7788-31</td><td></td></tr><tr><td>ET455/50m, DAPI-ET Emission Filter</td><td>Chroma Technology Corp.</td><td>ET455/50m</td><td></td></tr><tr><td>ET525/36m Emission Filter</td><td>Chroma Technology Corp.</td><td>ET525/36m</td><td></td></tr><tr><td>ET605/70m Emission Filter</td><td>Chroma Technology Corp.</td><td>ET605/70m</td><td></td></tr><tr><td>ET700/75m Emission Filter</td><td>Chroma Technology Corp.</td><td>ET700/75m</td><td></td></tr><tr><td>Bovine Serum Albumin Fraction V</td><td>RPI Products</td><td>A30075-100.0</td><td></td></tr><tr><td>Clara cooled charge-coupled device camera</td><td>Andor Technology</td><td>77026010</td><td></td></tr><tr><td>Cy3 donkey anti-rabbit</td><td>Jackson ImmunoResearch</td><td>711-165-152</td><td></td></tr><tr><td>DPBS</td><td>Thermo Fisher Scientific</td><td>14190250</td><td></td></tr><tr><td>Dyelight 649 goat anti-rat</td><td>Jackson ImmunoResearch</td><td>112-496-068</td><td></td></tr><tr><td>EGM-MV SingleQuot Supply &amp;amp; Growth Factors (contents shown below)</td><td>Lonza</td><td>CC-4143</td><td></td></tr><tr><td>Fetal bovine serum 25 ml</td><td></td><td></td><td></td></tr><tr><td>Bovine brain extract 2.0 ml</td><td></td><td></td><td></td></tr><tr><td>Gentamycin sulfate 0.5 ml</td><td></td><td></td><td></td></tr><tr><td>Epidermal growth factor 0.5 ml</td><td></td><td></td><td></td></tr><tr><td>Hydrocortisone 0.5 ml</td><td></td><td></td><td></td></tr><tr><td>Penicillin/Streptomycin (10,000 IU Pen/ml, 10,000 &amp;#181;g Strep/ml)&amp;nbsp; 10.0 ml</td><td></td><td></td><td></td></tr><tr><td>EGTA</td><td>Sigma-Aldrich</td><td>E3884-100G</td><td></td></tr><tr><td>Electronic shutter</td><td>Sutter instrument</td><td>77016099</td><td></td></tr><tr><td>Fibronectin</td><td>EMD Millipore</td><td>FC010</td><td></td></tr><tr><td>Glutaraldehyde</td><td>Thermo Fisher Scientific</td><td>S80026</td><td></td></tr><tr><td>HALT Protease inhibitor</td><td>Thermo Fisher Scientific</td><td>PI-78425</td><td></td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>H4034</td><td></td></tr><tr><td>HRP goat anti-mouse</td><td>Jackson ImmunoResearch</td><td>115-036-062</td><td></td></tr><tr><td>HRP goat anti-rabbit</td><td>Jackson ImmunoResearch</td><td>111-036-045</td><td></td></tr><tr><td>Immuno-Blot PVDF Membrane</td><td>Bio-Rad</td><td>162-0174</td><td></td></tr><tr><td>Lanolin</td><td>Thermo Fisher Scientific</td><td>8006-54-0</td><td></td></tr><tr><td>Magnesium sulfate</td><td>Sigma-Aldrich</td><td>M2643-500G</td><td></td></tr><tr><td>Microscope</td><td>Nikon TiE</td><td>MEA53100</td><td></td></tr><tr><td>Motorized stage</td><td>ASI Technologies</td><td>MS-2000</td><td></td></tr><tr><td>Mounting medium</td><td>Dako</td><td>CS70330-2</td><td></td></tr><tr><td>Mouse anti-Aurora A</td><td>Abcam</td><td>ab130156</td><td></td></tr><tr><td>Mouse anti-GAPDH</td><td>Abcam</td><td>ab9484</td><td></td></tr><tr><td>Mouse anti-MCAK</td><td>Abcam</td><td>ab50778</td><td></td></tr><tr><td>Mouse anti-tubulin (DM1A)</td><td>Cell Signaling Technology</td><td>3873S</td><td></td></tr><tr><td>Nail Polish</td><td>Electron Microscopy Sciences</td><td>72180</td><td></td></tr><tr><td>NIS Elements software</td><td>Nikon</td><td>MQS310000</td><td></td></tr><tr><td>Nucleofector Kit V</td><td>Lonza</td><td>VACA-1003</td><td>Contains the electroporation buffer</td></tr><tr><td>Paraffin</td><td>Thermo Fisher Scientific</td><td>P31-500</td><td></td></tr><tr><td>Penicillin-Streptomycin (10,000 IU Pen/ml, 10,000 &amp;#181;g Strep/ml)</td><td>MP Biomedicals</td><td>91670249</td><td></td></tr><tr><td>Petroleum jelly</td><td>Dow Corning</td><td>2021854-1113</td><td></td></tr><tr><td>Pipes</td><td>Sigma-Aldrich</td><td>P6757-100G</td><td></td></tr><tr><td>plusTipTracker</td><td></td><td></td><td></td></tr><tr><td>Rabbit anti-GFP</td><td>Abcam</td><td>ab290</td><td></td></tr><tr><td>Rabbit anti&amp;ndash;pT288&amp;ndash;Aurora A</td><td>Abcam</td><td>ab18318</td><td></td></tr><tr><td>Rat anti&amp;ndash;&amp;alpha;-tubulin</td><td>AbD Serotec</td><td>MCA78G</td><td></td></tr><tr><td>Sodium borohydride</td><td>Thermo Fisher Scientific</td><td>678-10</td><td></td></tr><tr><td>Spinning disk&amp;nbsp;</td><td>Andor Technology</td><td>Yokagawa CSU-X1</td><td></td></tr><tr><td>Sulfo-SANPAH</td><td>Thermo Fisher Scientific</td><td>PI-22589</td><td></td></tr><tr><td>SuperSignal West Pico Chemiluminescent Substrate</td><td>Thermo Fisher Scientific</td><td>34080</td><td></td></tr><tr><td>Triton X-100</td><td>Thermo Fisher Scientific</td><td>BP151-500</td><td></td></tr><tr><td>Trypsin</td><td>Corning</td><td>MT25053CI</td><td></td></tr><tr><td>Tween-20</td><td>Thermo Fisher Scientific</td><td>BP337</td><td></td></tr></tbody></table>

high-resolution time-lapse imaging and automated analysis of microtubule dynamics in living human umbilical vein endothelial cells

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes within individual endothelial cells (ECs). These processes are critical for homeostatic maintenance such as wound healing, and are also crucial in promoting tumor growth and metastasis. EC morphology is defined by the organization of the cytoskeleton, a tightly regulated system of actin and microtubule (MT) dynamics that is known to control EC branching, polarity and directional migration, essential components of angiogenesis. To study MT dynamics, we used high-resolution fluorescence microscopy coupled with computational image analysis of fluorescently-labeled MT plus-ends to investigate MT growth dynamics and the regulation of EC branching morphology and directional migration. Time-lapse imaging of living Human Umbilical Vein Endothelial Cells (HUVECs) was performed following transfection with fluorescently-labeled MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics. PlusTipTracker software was used to track EB3-labeled MT plus ends in order to measure MT growth speeds and MT growth lifetimes in time-lapse images. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration.

HUVEC culture for live-cell microscopy 
Live-cell imaging of HUVECs expressing fluorescent proteins requires that HUVECs are maintained in an environment that is suitable for normal cell growth and maintenance, as well as normal protein expression and function. Figure 1 shows a schematic depiction of the protocol used to prepare a buffered and closed system that maintains optimized optical characteristics for collecting time-lapse images of live cells. Thin strips of double-sided tape are placed onto a glass slide (Figure 1A) and then the coverslip containing the cultured HUVECs is inverted on top of the tape such that the cells are facing the slide (Figure 1B). In this way, the tape provides a small gap between the two glass surfaces, and creates a space in which the cells can be bathed in appropriate cell culture media (Figure 1C; see Protocol 4, above). In a final step, the glass coverslip is sealed to the glass slide using a petroleum jelly: lanolin: paraffin wax (1:1:1 mixture, VALAP; Figure 1D) to establish a closed system. This cell culture approach was used for both the shorter time sequence imaging of EB3 comets and for the longer time sequence branching and migration experiments. Additionally, the wax-like consistency of VALAP allows for easy removal from the glass slide and coverslip at the end of an imaging experiment such that supplemental approaches including fixation and immunolabelling (see Protocol 8), can be performed on the same coverslip and cell sample that was visualized using the live-cell imaging approach.
<img alt="Figure 1" src="/files/ftp_upload/54265/54265fig1.jpg" /> 
Figure 1: Methodology of HUVEC culture for live-cell microscopic imaging. (A) Cut 2 pieces of double-sided tape into 0.65 cm x 2.50 cm rectangular strips and secure one piece of double-sided tape horizontally along the top edge of the slide, and the other piece, along the bottom edge of the slide. It is important to allow a small amount of space between the tape and the slide edge for sealing the chamber as described in (D). (B) Using forceps remove the coverslip containing HUVECs and invert the coverslip (cell side down) on top of the pieces of tape. (C) Using a micropipette, add 40 &#956;l of imaging media (complete endothelial basal medium supplemented with 25 &#956;M HEPES) between the coverslip and the glass slide. Ensure that the space between the coverslip and slide is completely filled with imaging media and avoid bubbles. Use an aspirator along the edge of the coverslip to remove excess media, if necessary. (D) Use warm VALAP (1:1:1 vaseline : lanolin : paraffin) to seal around the edges of the coverslip. Check for imaging media leakages by pushing around gently on the coverslip. Use additional VALAP to remedy. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54265/54265fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Automated tracking of fluorescently-labeled EB3 allows for comprehensive analysis of whole cell and regional MT growth dynamics. The collection of clear, distinct, and highly time and space-resolved microscopic images of the EB3 protein is essential to the analysis of EB3 movements within the cell. HUVEC expression of fluorescently-labeled EB3 varies widely from cell to cell and fluorescence intensity typically increases within the same cell over a period of hr. Thus, it is important to consider and monitor these changes in expression profile in order to identify potential effects of increased EB expression on MT growth dynamics. This is best accomplished by evaluating grey-scale fluorescence intensity profiles for each imaged cell, and then restricting data analysis to those cells that display a defined range of grey-scale expression profiles. Initial evaluation of expression profile data sets is critical, and should be compared with MT growth dynamics data that is obtained by plusTipTracker software for each cell. Expression profiles can be plotted against MT growth speed and MT growth lifetime for each cell within an experimental group, in order to determine the desired range of grey-scale expression and to identify potential outliers.
Figure 2 highlights a HUVEC cultured on a fibronectin substrate (see Protocol 2.1, above) and expressing near optimal levels of mApple-labeled EB3. Fluorescent EB3 comets are shown in a time-lapse series of images over a period of 12 sec (Figure 2A). Analysis of EB3 comets using plusTipTracker software involves an automated, frame-by-frame EB3 comet detection (green and yellow dots, Figure 2B). Comets detected within each frame are then linked, dependent upon their change in position from frame-to-frame, to generate EB3 tracks and a visual EB3 track overlay (Figure 2C-F). Track overlays are color-coded and color distinctions can be determined by parameters entered by the user. Commonly, these distinctions use the global mean of MT growth speed and MT growth lifetime5,31 and thereby split the data into four groups: slow and short-lived (red), slow and long-lived (yellow), fast and short-lived (green), and fast and long-lived MT growth (blue; Figure 2G). PlusTipTracker-mediated tracking of EB3-labeled MT growth dynamics also enables user-defined Regions of Interest (ROIs). PlusTipTracker extracts the MT growth track data from within the user-defined ROI (Figure 2A, B and F), thereby allowing for comparisons of MT growth dynamics within different regions of the same cell.
<img alt="Figure 2" src="/files/ftp_upload/54265/54265fig2.jpg" /> 
Figure 2: EB3 comet detection, tracking, and plusTipTracker data output. (A) time-lapse movie of a HUVEC expressing mApple EB3. Left panel shows a whole cell view and whole cell mask (white line) demarking the cell boundaries. Red box represents the zoomed region shown in the time-lapse images (right panels). Right panels show raw grey-scale time-lapse images of EB3 comets in successive imaging frames (t = 0 sec - t = 12 sec). (B) plusTipTracker detection of EB3 comets from the images shown in (A). Left panel shows a whole cell view of detected comets (yellow dots). Red box represents the zoomed region shown in the time-lapse images (right panels). Right panels highlight five detected EB3 comets (red arrows) and trace those five comets over the 12 sec imaging period. Green dots represent detected EB3 comets that did not exist (or were not detected) in the previous time interval. (C) Maximum intensity projection of 61 frames (2 min movie) of the EB3 comets shown in (A). (D) plusTipTracker MT track overlay of the same 61 frames (2 min movie) shown in (C). (E) Zoom of maximum intensity projection (red box in C). (F) Zoom of plusTipTracker MT track overlay of maximum intensity projection (red box in D). (G) Color-coded legend for track overlays shown in (D) and (F). Designation as slow, fast, short-lived or long-lived is based on the mean value for all growth tracks measured in a group of ten mApple-EB3 expressing HUVECs. For easier visualization, track overlays in (D) and (F) are generated using inverted pixel intensities of the cell image shown in (A). Scale bars = 20 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54265/54265fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
HUVEC branching and migration patterns are sensitive to mechano-physical signals from the ECM.
HUVECs are well known to respond to various types of signaling cues and, in doing so, to undergo morphological alterations that dictate an appropriate cellular response to the particular signaling cue(s)36,40-42. HUVECs cultured on fibronectin coated glass substrates (Figure 3A), or on stiff 2-dimensional polyacrylamide ECMs (55 kPa) typically display a round or elongated morphology with few distinct branched protrusions (Figure 3B). However, when cultured on very soft 2-dimensional polyacrylamide ECMs (0.7 kPa), HUVECs display a highly branched morphology that is known to result from compliance-induced downregulation of myosin-II contractility4,5,31 (Figure 3C). Thus, HUVECs regulate their cellular morphology through ECM mechanosensing, and this is accomplished through localized modulation of MT dynamics and acto-myosin contractility.
Because HUVECs display a wide variety of morphological attributes, in order to measure HUVEC branching it is critical to first define what a branched protrusion is, and then to define how a branched protrusion will be measured objectively. One technique uses a four-step approach by which a cell boundary or &#34;mask&#34; is generated by hand tracing the edges of the cell using an image of a cell expressing a fluorescent volume marker to define the cell edges (Figure 3D). Following mask generation, branch origins are defined by locating greatest angle of curvature between the branched protrusion and the cell body. A line is drawn to demark the &#34;branch origin&#34; (purple angled lines; Figure 3E). Branch number is measured simply by counting the number of branches with a defined branch origin (Figure 3F). Branch length is measured as the distance from the branch origin to the distal branch tip (Figure 3G). To avoid inclusion of small lamellipodial projections from data analysis, additional criterion require that branches must be longer (&#34;branch length&#34;) than they are wide (at the &#34;branch origin&#34;), and that branches must be at least ten microns in length5,31.
<img alt="Figure 3" src="/files/ftp_upload/54265/54265fig3.jpg" /> 
Figure 3: Analysis of HUVEC branching morphology. (A-C) DIC image examples of HUVECs cultured on fibronectin coated glass ECMs (A) stiff (55 kPa) polyacrylamide ECMs (B) and soft (0.7 kPa) polyacrylamide ECMs (C). Branching is dramatically increased in HUVECs cultured on 0.7 kPa polyacrylamide ECMs (C) as compared to stiffer ECMs (A, B). (D). DIC microscopic image (left image) of a transfected HUVEC expressing a mApple-C1 cDNA construct (a cell volume marker; right image). (E) Define the branch origin by locating the greatest angle of curvature (i.e. the position where the membrane displays the greatest curvature; purple-colored lines) on either side of the branch and then connect the angles with a straight line (blue lines). (F) Identify and selecting branched protrusions that extend &#62;10 &#956;m from the &#34;branch origin&#34; (defined in E). Count the number of branched protrusions to obtain the &#34;branch number.&#34; (G) Measure the distance from the center of the &#34;branch origin&#34; to the most distal point of the branch using the line tool in the annotations and measurements application of NIS Elements Software to obtain the &#34;branch length.&#34; Scale bars = 20 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54265/54265fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
In addition to sensing ECM compliance, HUVECs also sense and respond to the elimination of adjacent cellular contacts as induced by scratch wounding. In the migration assay, HUVECs cultured in a monolayer are subjected to a scratch wound and migration characteristics are measured (see Protocol 9, above). After induction of the wound by scratching the monolayer, wound-edge HUVECs polarize by developing a protrusive leading edge that extends into the wound (Figure 4A, t = 0 hr). Over a time course of 10-12 hr, polarized wound-edge HUVECs migrate into the wound area and fill the wound, establishing a new confluent monolayer (Figure 4A, t = 11 hr). As with HUVEC branching, current evidence suggests that polarization and migration is critically dependent upon the coordinated re-organization of the MT and actin cytoskeletons43-49. This is achieved, in part, through localized inhibition of MCAK-induced MT disassembly at the leading edge, but not at the trailing edge of wound-edge HUVECs31. The sensitivity of MT dynamics to wound-induced polarization is initiated by the Rac1 GTPase, which targets numerous downstream regulator proteins, including Aurora-A kinase that phosphorylates and inhibits MCAK at the leading edge, and other kinases that are capable of inhibiting or activating MAPs to locally modulate MT growth and shrinkage18,31,36,48,50-53. Extensive experimental evidence supports the notion that coordination of leading edge protrusion and trailing edge contraction is governed through cycles of Rac1 activation to promote MT assembly at the leading edge and RhoA activation to acto-myosin contractility at the trailing edge, thereby driving directed motility9,41,51,52,54-58.
To date, there is not an established methodology for performing automated tracking of cell migration. This is a result of difficulty in computationally analyzing cell-ECM boundaries that are consistently changing as wound-edge cells polarize and migrate. While many software programs are capable of tracking fluorescently labeled cells59-64, the population of fluorescently labeled HUVECs in wound-edge migration experiments is approximately 30-40% of the total HUVEC population, and therefore the majority of the wound-edge cells must be visualized by phase-contrast or DIC imaging. Additionally, it is important when measuring wound-edge migration to evaluate fluorescent and non-fluorescent cells within the same wound in order to identify specific effects of expression constructs both within experimental study groups and between experimental study groups. Currently, the best approach to measure HUVEC migration is to use a hand tracking approach in which a wound-edge cell is first identified and then the nucleus and nucleolus are identified (Figure 4B). The nucleolus is used as the position determinant for cell tracking due to its high light scattering density and relatively small size. These two features of the nucleolus make it easier to identify and reduce measurement error by limiting the area in which the user can place individual track positions. HUVEC migration tracks are then generated by clicking on the nucleolus in successive time-lapse image frames. Finally, migration velocity and migration distance to origin are measured and analyzed for control and experimental cell groups (Figure 4B).
<img alt="Figure 4" src="/files/ftp_upload/54265/54265fig4.jpg" /> 
Figure 4: Analysis of wound-edge HUVECs and migration tracking. (A) 20X phase-contrast images of an 11 hr time-lapse imaging series of scratch wound-edge HUVECs (a.k.a. &#34;Migration Assay&#34;). The wound and direction of migration are indicated in panel t = 0 hr. HUVECs are shown to cross the wound-edge and advance into the wound until a monolayer of cells is formed (t = 3.5 hr - t = 11 hr). (B) Analysis of migration assay time lapse imaging (as shown in A) requires first the identification of a trackable wound-edge HUVEC based on cell location (i.e. cells physically at the edge of the wound) and expression profile (i.e., select a green wound edge cell if the experiment involves expression of GFP). Using the phase contrast image, identify the nucleus and the nucleolus. Hand track the nucleolus in successive time-lapse images using the annotations and measurements application in NIS Elements software to determine migration velocity and distance to origin. Scale bars = 100 &#956;m&#160; (A), 20 &#956;m (B). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54265/54265fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells at JoVE.com

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed.

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes ...

high-resolution-time-lapse-imaging-automated-analysis-microtubule

Nanyang Technological University

Research

JoVE Journal

Biology

11.0K Views.  University of the Sciences in Philadelphia. Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed. 

Video: High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

Isolation of Human Umbilical Vein Endothelial Cells (HUVEC)

Angiogenesis is a complex multi-step process, where in response to angiogenic stimuli, new vessels are created from the existing vasculature. These steps include: degradation of the basement membrane, proliferation and migration (sprouting) of endothelial cells (EC) into the extracellular matrix, alignment of EC into cords, lumen formation, anastomosis, and formation of a new basement membrane. Many in vitro assays have been developed to study this process, but most only mimic certain stages of angiogenesis, and morphologically the vessels often do not resemble vessels in vivo. Here we demonstrate an optimized in vitro angiogenesis assay that utilizes human umbilical vein EC and fibroblasts. This model recapitulates all of the key early stages of angiogenesis, and importantly the vessels display patent intercellular lumens surrounded by polarized EC. Vessels can be easily observed by phase-contrast and time-lapse microscopy, and recovered in pure form for downstream applications.

Isolation of Human Umbilical Vein Endothelial Cells HUVEC

This video protocol illustrates the isolation and culture of human umbilical vein endothelial cells (HUVEC) from human umbilical cord. Once isolated these cells can be used for in vitro angiogenesis assays like the Optimized Fibrin Gel Bead Assay also demonstrated by the Hughes lab.

Angiogenesis is a complex multi-step process, where in response to angiogenic stimuli, new vessels are created from the existing vasculature. These steps ...

Isolation of Human Umbilical Vein Endothelial Cells and Their Use in the Study of Neutrophil Transmigration Under Flow Conditions

Neutrophils are the most abundant type of white blood cell. They form an essential part of the innate immune system1. During acute inflammation, neutrophils are the first inflammatory cells to migrate to the site of injury. Recruitment of neutrophils to an injury site is a stepwise process that includes first, dilation of blood vessels to increase blood flow; second, microvascular structural changes and escape of plasma proteins from the bloodstream; third, rolling, adhesion and transmigration of the neutrophil across the endothelium; and fourth accumulation of neutrophils at the site of injury2,3. A wide array of in vivo and in vitro methods has evolved to enable the study of these processes4. This method focuses on neutrophil transmigration across human endothelial cells.
One popular method for examining the molecular processes involved in neutrophil transmigration utilizes human neutrophils interacting with primary human umbilical vein endothelial cells (HUVEC)5. Neutrophil isolation has been described visually elsewhere6; thus this article will show the method for isolation of HUVEC. Once isolated and grown to confluence, endothelial cells are activated resulting in the upregulation of adhesion and activation molecules. For example, activation of endothelial cells with cytokines like TNF-&alpha; results in increased E-selectin and IL-8 expression7. E-selectin mediates capture and rolling of neutrophils and IL-8 mediates activation and firm adhesion of neutrophils. After adhesion neutrophils transmigrate. Transmigration can occur paracellularly (through endothelial cell junctions) or transcellularly (through the endothelial cell itself). In most cases, these interactions occur under flow conditions found in the vasculature7,8.
The parallel plate flow chamber is a widely used system that mimics the hydrodynamic shear stresses found in vivo and enables the study of neutrophil recruitment under flow condition in vitro9,10. Several companies produce parallel plate flow chambers and each have advantages and disadvantages. If fluorescent imaging is needed, glass or an optically similar polymer needs to be used. Endothelial cells do not grow well on glass.
Here we present an easy and rapid method for phase-contrast, DIC and fluorescent imaging of neutrophil transmigration using a low volume ibidi channel slide made of a polymer that supports the rapid adhesion and growth of human endothelial cells and has optical qualities that are comparable to glass. In this method, endothelial cells were grown and stimulated in an ibidi &mu;slide. Neutrophils were introduced under flow conditions and transmigration was assessed. Fluorescent imaging of the junctions enabled real-time determination of the extent of paracellular versus transcellular transmigration.

This article first describes a procedure for isolating human endothelial cells from umbilical veins and then shows how to use these cells to examine neutrophil transmigration under flow conditions. By using a low-volume flow chamber made from a polymer with the optical characteristics of glass, live-cell fluorescent imaging of rare cell populations is also possible.

Neutrophils are the most abundant type of white blood cell. They form an essential part of the innate immune system1. During acute inflammation, ...

Immunology and Infection

Real-time Imaging of Endothelial Cell-cell Junctions During Neutrophil Transmigration Under Physiological Flow

During inflammation, leukocytes leave the circulation and cross the endothelium to fight invading pathogens in underlying tissues. This process is known as leukocyte transendothelial migration. Two routes for leukocytes to cross the endothelial monolayer have been described: the paracellular route, i.e., through the cell-cell junctions and the transcellular route, i.e., through the endothelial cell body. However, it has been technically difficult to discriminate between the para- and transcellular route. We developed a simple in vitro assay to study the distribution of endogenous VE-cadherin and PECAM-1 during neutrophil transendothelial migration under physiological flow conditions. Prior to neutrophil perfusion, endothelial cells were briefly treated with fluorescently-labeled antibodies against VE-cadherin and PECAM-1. These antibodies did not interfere with the function of both proteins, as was determined by electrical cell-substrate impedance sensing and FRAP measurements. Using this assay, we were able to follow the distribution of endogenous VE-cadherin and PECAM-1 during transendothelial migration under flow conditions and discriminate between the para- and transcellular migration routes of the leukocytes across the endothelium.

Leukocytes cross the endothelial monolayer using the paracellular or the transcellular route. We developed a simple assay to follow the distribution of endogenous junctional VE-cadherin and PECAM-1 during leukocyte transendothelial migration under physiological flow to discriminate between the two transmigration routes.

During inflammation, leukocytes leave the circulation and cross the endothelium to fight invading pathogens in underlying tissues. This process is known ...

Micropatterning and Assembly of 3D Microvessels

In vitro platforms to study endothelial cells and vascular biology are largely limited to 2D endothelial cell culture, flow chambers with polymer or glass based substrates, and hydrogel-based tube formation assays. These assays, while informative, do not recapitulate lumen geometry, proper extracellular matrix, and multi-cellular proximity, which play key roles in modulating vascular function. This manuscript describes an injection molding method to generate engineered vessels with diameters on the order of 100 &#181;m. Microvessels are fabricated by seeding endothelial cells in a microfluidic channel embedded within a native type I collagen hydrogel. By incorporating parenchymal cells within the collagen matrix prior to channel formation, specific tissue microenvironments can be modeled and studied. Additional modulations of hydrodynamic properties and media composition allow for control of complex vascular function within the desired microenvironment. This platform allows for the study of perivascular cell recruitment, blood-endothelium interactions, flow response, and tissue-microvascular interactions. Engineered microvessels offer the ability to isolate the influence from individual components of a vascular niche and precisely control its chemical, mechanical, and biological properties to study vascular biology in both health and disease.

This manuscript presents an injection molding method to engineer microvessels that recapitulate physiological properties of endothelium. The microfluidic-based process creates patent 3D vascular networks with tailorable conditions, such as flow, cellular composition, geometry, and biochemical gradients. The fabrication process and examples of potential applications are described.

In vitro platforms to study endothelial cells and vascular biology are largely limited to 2D endothelial cell culture, flow chambers with polymer or glass ...

Bioengineering

Extended Time-lapse Intravital Imaging of Real-time Multicellular Dynamics in the Tumor Microenvironment

In the tumor microenvironment, host stromal cells interact with tumor cells to promote tumor progression, angiogenesis, tumor cell dissemination and metastasis. Multicellular interactions in the tumor microenvironment can lead to transient events including directional tumor cell motility and vascular permeability. Quantification of tumor vascular permeability has frequently used end-point experiments to measure extravasation of vascular dyes. However, due to the transient nature of multicellular interactions and vascular permeability, the kinetics of these dynamic events cannot be discerned. By labeling cells and vasculature with injectable dyes or fluorescent proteins, high-resolution time-lapse intravital microscopy has allowed the direct, real-time visualization of transient events in the tumor microenvironment. Here we describe a method for using multiphoton microscopy to perform extended intravital imaging in live mice to directly visualize multicellular dynamics in the tumor microenvironment. This method details cellular labeling strategies, the surgical preparation of a mammary skin flap, the administration of injectable dyes or proteins by tail vein catheter and the acquisition of time-lapse images. The time-lapse sequences obtained from this method facilitate the visualization and quantitation of the kinetics of cellular events of motility and vascular permeability in the tumor microenvironment.

This protocol describes the use of multiphoton microscopy to perform extended time-lapse imaging of multicellular interactions in real time, in vivo at single cell resolution.

In the tumor microenvironment, host stromal cells interact with tumor cells to promote tumor progression, angiogenesis, tumor cell dissemination and ...

Medicine

Spatial Temporal Analysis of Fieldwise Flow in Microvasculature

Changes in blood flow velocity and distribution are vital in maintaining tissue and organ perfusion in response to varying cellular needs. Further, appearance of defects in microcirculation can be a primary indicator in the development of multiple pathologies. Advances in optical imaging have made intravital microscopy (IVM) a practical approach, permitting imaging at the cellular and subcellular level in live animals at high-speed over time. Yet, despite the importance of maintaining adequate tissue perfusion, spatial and temporal variability in capillary flow is seldom documented. In the standard approach, a small number of capillary segments are chosen for imaging over a limited time. To comprehensively quantify capillary flow in an unbiased way we developed Spatial Temporal Analysis of Fieldwise Flow (STAFF), a macro for FIJI open-source&#160;image analysis software. Using high-speed image sequences of full fields of blood flow within capillaries, STAFF produces&#160;images that represent motion over time called kymographs for every time interval for every vascular segment. From the kymographs STAFF calculates velocities from the distance that red blood cells move over time, and outputs the velocity data as a sequence of color-coded spatial maps for visualization and tabular output for quantitative analyses. In normal mouse livers, STAFF analyses quantified profound differences in flow velocity between pericentral and periportal regions within lobules. Even more unexpected are the differences in flow velocity seen between sinusoids&#160;that are side by side and fluctuations seen within individual vascular segments over seconds. STAFF is a powerful new tool capable of providing novel insights by enabling measurement of the complex spatiotemporal dynamics of capillary flow.

To quantify microvascular flow from high speed capillary flow image sequences, we developed STAFF (Spatial Temporal Analysis of Fieldwise Flow) software. Across the full image field and over time, STAFF evaluates flow velocities and generates a sequence of color-coded spatial maps for visualization and tabular output for quantitative analyses.

Changes in blood flow velocity and distribution are vital in maintaining tissue and organ perfusion in response to varying cellular needs. Further, ...

Real-Time Intravital Multiphoton Microscopy to Visualize Focused Ultrasound and Microbubble Treatments to Increase Blood-Brain Barrier Permeability

The blood-brain barrier (BBB) is a key challenge for the successful delivery of drugs to the brain. Ultrasound exposure in the presence of microbubbles has emerged as an effective method to transiently and locally increase the permeability of the BBB, facilitating para- and transcellular transport of drugs across the BBB. Imaging the vasculature during ultrasound-microbubble treatment will provide valuable and novel insights on the mechanisms and dynamics of ultrasound-microbubble treatments in the brain.
Here, we present an experimental procedure for intravital multiphoton microscopy using a cranial window aligned with a ring transducer and a 20x objective lens. This set-up enables high spatial and temporal resolution imaging of the brain during ultrasound-microbubble treatments. Optical access to the brain is obtained via an open-skull cranial window. Briefly, a 3-4 mm diameter piece of the skull is removed, and the exposed area of the brain is sealed with a glass coverslip. A 0.82 MHz ring transducer, which is attached to a second glass coverslip, is mounted on top. Agarose (1% w/v) is used between the coverslip of the transducer and the coverslip covering the cranial window to prevent air bubbles, which impede ultrasound propagation. When sterile surgery procedures and anti-inflammatory measures are taken, ultrasound-microbubble treatments and imaging sessions can be performed repeatedly over several weeks. Fluorescent dextran conjugates are injected intravenously to visualize the vasculature and quantify ultrasound-microbubble induced effects (e.g., leakage kinetics, vascular changes). This paper describes the cranial window placement, ring transducer placement, imaging procedure, common troubleshooting steps, as well as advantages and limitations of the method.

This protocol describes the surgical and technical procedures that enable real-time in vivo multiphoton fluorescence imaging of the rodent brain during focused ultrasound and microbubble treatments to increase blood-brain barrier permeability.

The blood-brain barrier (BBB) is a key challenge for the successful delivery of drugs to the brain. Ultrasound exposure in the presence of microbubbles ...

Microtubule Plus-End Dynamics Visualization in Huntington's Disease Model based on Human Primary Skin Fibroblasts

Transfection with a fluorescently labeled marker protein of interest in combination with time-lapse video microscopy is a classic method of studying the dynamic properties of the cytoskeleton. This protocol offers a technique for human primary fibroblast transfection, which can be difficult because of the specifics of primary cell cultivation conditions. Additionally, cytoskeleton dynamic property maintenance requires a low level of transfection to obtain a good signal-to-noise ratio without causing microtubule stabilization. It is important to take measures to protect the cells from light-induced stress and fluorescent dye fading. In the course of our work, we tested different transfection methods and protocols as well as different vectors to select the best combination of conditions suitable for human primary fibroblast studies. We analyzed the resulting time-lapse videos and calculated microtubule dynamics using ImageJ. The dynamics of microtubules' plus-ends in the different cell parts are not similar, so we divided the analysis into subgroups - the centrosome region, the lamella, and the tail of fibroblasts. Notably, this protocol can be used for in vitro analysis of cytoskeleton dynamics in patient samples, enabling the next step towards understanding the dynamics of the various disease development.

This protocol is dedicated to the microtubule plus-end visualization by EB3 protein transfection to study their dynamic properties in primary cell culture. The protocol was implemented on human primary skin fibroblasts obtained from Huntington's disease patients.

Transfection with a fluorescently labeled marker protein of interest in combination with time-lapse video microscopy is a classic method of studying the ...

Automated Analysis of Dynamic Ca2+ Signals in Image Sequences

Intracellular Ca2+ signals are commonly studied with fluorescent Ca2+ indicator dyes and microscopy techniques. However, quantitative analysis of Ca2+ imaging data is time consuming and subject to bias. Automated signal analysis algorithms based on region of interest (ROI) detection have been implemented for one-dimensional line scan measurements, but there is no current algorithm which integrates optimized identification and analysis of ROIs in two-dimensional image sequences. Here an algorithm for rapid acquisition and analysis of ROIs in image sequences is described. It utilizes ellipses fit to noise filtered signals in order to determine optimal ROI placement, and computes Ca2+ signal parameters of amplitude, duration and spatial spread. This algorithm was implemented as a freely available plugin for ImageJ (NIH) software. Together with analysis scripts written for the open source statistical processing software R, this approach provides a high-capacity pipeline for performing quick statistical analysis of experimental output. The authors suggest that use of this analysis protocol will lead to a more complete and unbiased characterization of physiologic Ca2+ signaling.

Automated Analysis of Dynamic Ca2+ Signals in Image Sequences

Here a novel region of interest analysis protocol based on sorting best-fit ellipses assigned to regions of positive signal within two-dimensional time lapse image sequences is demonstrated. This algorithm may enable investigators to comprehensively analyze physiological Ca2+ signals with minimal user input and bias.

Intracellular Ca2+ signals are commonly studied with fluorescent Ca2+ indicator dyes and microscopy techniques. However, quantitative analysis of Ca2+ ...

Development and Characterization of In Vitro Microvessel Network and Quantitative Measurements of Endothelial [Ca2+]i and Nitric Oxide Production

Endothelial cells (ECs) lining the blood vessel walls in vivo are constantly exposed to flow, but cultured ECs are often grown under static conditions and exhibit a pro-inflammatory phenotype. Although the development of microfluidic devices has been embraced by engineers over two decades, their biological applications remain limited. A more physiologically relevant in vitro microvessel model validated by biological applications is important to advance the field and bridge the gaps between in vivo and in vitro studies. Here, we present detailed procedures for the development of cultured microvessel network using a microfluidic device with a long-term perfusion capability. We also demonstrate its applications for quantitative measurements of agonist-induced changes in EC [Ca2+]i and nitric oxide (NO) production in real time using confocal and conventional fluorescence microscopy. The formed microvessel network with continuous perfusion showed well-developed junctions between ECs. VE-cadherin distribution was closer to that observed in intact microvessels than statically cultured EC monolayers. ATP-induced transient increases in EC [Ca2+]i and NO production were quantitatively measured at individual cell levels, which validated the functionality of the cultured microvessels. This microfluidic device allows ECs to grow under a well-controlled, physiologically relevant flow, which makes the cell culture environment closer to in vivo than that in the conventional, static 2D cultures. The microchannel network design is highly versatile, and the fabrication process is simple and repeatable. The device can be easily integrated to the confocal or conventional microscopic system enabling high resolution imaging. Most importantly, because the cultured microvessel network can be formed by primary human ECs, this approach will serve as a useful tool to investigate how pathologically altered blood components from patient samples affect human ECs and provide insight into clinical issues. It also can be developed as a platform for drug screening.

Development and Characterization of In Vitro Microvessel Network and Quantitative Measurements of Endothelial [Ca2+]i and Nitric Oxide Production

Primary human umbilical vein endothelial cells (HUVECs) were grown to confluence within a microfluidic network device. The endothelial cell junction and F-actin distributions were illustrated and the changes in intracellular calcium concentration and nitric oxide production in response to adenosine triphosphate (ATP) were quantified in real-time at individual cell levels.

Endothelial cells (ECs) lining the blood vessel walls in vivo are constantly exposed to flow, but cultured ECs are often grown under static conditions and ...

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

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